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Endogenous beta-galactosidase activity in the larval, pupal, and adult stages of the fruit fly, Drosophila melanogaster, indicates need for caution in lacZ fusion-gene studies.

Introduction

The glycosidase [Beta]-galactosidase is known to occur in a variety of microorganisms, plants, and animals (Wallenfels and Weil, 1972), and the fruit fly Drosophila melanogaster is no exception (Knipple and MacIntyre, 1984; Fuerst et al., 1987; Glaser et al., 1986). This enzyme, which cleaves the [Beta]-glycosidic bond between D-galactose and other sugar moieties, has been extensively studied in the prokaryote Escherichia coli, where it cleaves lactose into glucose and galactose.

With the advent of recombinant DNA technology, E. coli [Beta]-galactosidase and its structural gene, lacZ, have become very useful in molecular studies. Because lacZ has been sequenced and cloned, and produces an enzyme easily detectable with various chromogenic assays, it is used extensively as a reporter gene to trace the spatial and temporal expression of genes in transgenic organisms. Before it is inserted into another genome, the E. coli lacZ reporter gene is fused to the promoter, with or without additional sequences, of a gene whose expression is to be monitored. When this lacZ fusion gene becomes incorporated into the genome of an organism, its expression can be detected by assaying for E. coli [Beta]-galactosidase. Once the elegant P-element-mediated mutagenesis technique was developed for inserting genes into the genome of Drosophila melanogaster (for review see Spradling, 1986), the fruit fly became a perfect recipient for lacZ fusion genes, and transformed flies with fusion genes containing an E. coli lacZ reporter gene became common.

Typically in these transformed flies, tissues are assayed for the reporter gene product, E. coli [Beta]-galactosidase, by using the chromogen X-gal (5-bromo-4-chloro-3-indolyl-[Beta]-D-galactoside). Here, [Beta]-galactosidase hydrolyzes the [Beta]-glycosidic bond of X-gal and produces an insoluble blue stain of indolyl monomers. Because minute amounts of enzyme can be detected with the X-gal stain, it is an effective method for tagging fusion-gene expression. The lacZ fusion gene/X-gal system is an economical and rapid assay of tissue-specific gene activity in D. melanogaster, it is even more sensitive than immunocytochemical (Ghysen and O'Kane, 1989) or in situ hybridization techniques.

However, when assaying E. coli lacZ fusion-gene activity with X-gal, it is important to remember that these transformed fruit flies will express an endogenous [Beta]-galactosidase as well as the E. coli enzyme. Where the endogenous [Beta]-galactosidase is present, it will hydrolyze X-gal and will be indistinguishable from its ectopic counterpart - even when the pH of the staining buffer has been adjusted to levels appropriate for optimal E. coli [Beta]-galactosidase activity (Ashburner, 1989).

An endogenous [Beta]-galactosidase, [Beta]-galactosidase-1, has been isolated from adult D. melanogaster (Knipple and MacIntyre, 1984) and characterized as a glycoprotein homodimer with an optimal activity (hydrolysis of p-nitrophenyl-[Beta]-D-galactopyranoside) at a pH of 6.0 (Fuerst et al., 1987). Furthermore, three electrophoretically distinct minor forms of the enzyme have been found in other developmental stages of D. melanogaster (Knipple, 1983). Despite these findings, endogenous [Beta]-galactosidase expression has been thought to be negligible and has not been regarded as a problem in lacZ fusion-gene studies in D. melanogaster (see, for example, Lawrence, 1992, p. 52).

In this study, we show that endogenous [Beta]-galactosidase activity is not negligible. Using an X-gal assay identical to those employed to detect E. coli lacZ fusion-gene products, we identified the tissues that express endogenous [Beta]-galactosidase activity in the late third-instar larva, prepupa, pupa, and adult of wild-type D. melanogaster. Specifically, we assayed for [Beta]-galactosidase activity in active and nonactive third instar larvae: 15 metamorphic stages; and newly eclosed adults and those at age intervals of 1 to 2, 3 to 5, and 7 to 10 days. The resulting maps of expression not only provide a guide to endogenous activity for investigators using a lacZ reporter gene in a wild-type background, but also characterize the distribution and intensity of endogenous [Beta]-galactosidase activity in different life stages, and suggest that endogenous [Beta]-galactosidase is intrinsically interesting in view of its association with metamorphic remodeling and phagocytic activity.

Materials and Methods

Axenic cultures

To prevent background staining due to commensals or other microorganisms that might express [Beta]-galactosidase, we reared a wild-type Gailey strain of D. melanogaster (a Canton-S derivative, kindly supplied by Dr. Donald Galley of California State University at Hayward) under sterile conditions (modified from Roberts, 1986). Mature adult flies were placed on fresh cornmeal/molasses/agar/yeast/malt medium (with proprionic acid added as a mold inhibitor) for 3-6 hours. Eggs were collected, placed in vials of fine-mesh plankton net, and sterilized in a solution of 5% sodium hypochorite (full-strength Clorox). After the chorion had sloughed off (10-15 min) and hatched larvae (if any) were removed, the eggs were rinsed three times in sterile distilled water, 10 min each rinse. A sterile pasteur pipette was used to transfer eggs to autoclaved vials of axenic medium (1.5 g agar/7 g cornmeal/7 g yeast/1 ml proprionic acid to 100 ml water). All egg-handling operations were carried out in a sterile laminar flow hood, and culture vials were stored in the sterile hood until sampling time.

Collection of developmental stages

Late third-instar larvae were collected as they crawled out of the food medium and were sampled as two groups: actively crawling larvae and older nonactive larvae just prior to eversion of the spiracles. We chose these particular groups of late third-instar larvae because they are often collected for sampling of larval tissues and imaginal discs. The only visible difference between the two groups is crawling activity; however, changes in endogenous [Beta]-galactosidase activity may take place in the nonactive larvae as they approach the prepupal stage. To track the tissue-specific activity of [Beta]-galactosidase through metamorphosis, we collected both prepupal and pupal stages (P1-P15, Bainbridge and Bownes, 1981). To determine whether the enzyme activity in adults varied with age or sex, we collected male and female flies at intervals of 0.5-3 h, 12 days, 3-5 days, and 7-10 days after eclosion.

Whole mounts

Whole-mount preparations were dissected from larval, pupal, and adult stages and stained with X-gal. To determine whether staining varied among individual animals, we dissected and stained 10 animals from each of the following stages: active and inactive late third-instar larvae: prepupal stages P1-P3: and newly eclosed adults and adults aged 1 to 2 days, 3 to 5 days, and 7 to 10 days. Older pupal stages were too fragile to dissect; thus they were not stained as whole mounts.

Animals were placed under a Nikon SMZ-U dissecting microscope and dissected with the aid of microforceps, iridectomy scissors, and 0.1-mm insect pins. Tools, pins, and small plastic petri dishes with a Silgard (Dow Coming Corp.) base were disinfected in 95% ethanol, and tissues were kept wet with phosphate-buffered saline (PBS), pH 7.2, that had been sterilized through a Gelman acrodisc filter assembly (0.2-[[micro]meter] poresize). Adult flies were anesthetized on ice or with ether before dissection. Any fat-body tissue surrounding organs was removed because it hardened during fixation, making identification of structures difficult.

Cryostat sectioning

Frozen serial sections (transverse sections of larvae and sagittal sections of pupae from stages P1-P15) were cut and stained with X-gal. Before they were frozen, pupae were removed from the puparia: super glue gel "Quick Gel" (Loctite Corp.) was used to adhere the ventral surface of the puparium to a petri dish; after the glue hardened, microforceps were used to dissect the puparium away from the pupa. This technique allowed rapid removal and accurate staging of the delicate pupa without damage. With the puparium removed, the pupal cuticle freezes in direct contact with the mounting medium and is firmly embedded within the block, allowing high-quality cryostat sections to be cut. For sectioning, the organisms were mounted in OTC (Miles Inc.) or TBS (Triangle Biomedical Sciences) on stubs cooled with dry ice. When storage before sectioning was necessary, mounted specimens were removed from the stubs and stored in airtight plastic bags at -80 [degrees] C. We found that the resolution of X-gal staining was optimal in thick (20-[[micro]meter]) sections. Serial sections were cut on a Reichert-Jung 975C cryostat microtome at -18 [degrees] to -21 [degrees] C, collected in order on gelatin-coated slides, and air-dried for 1-3 h before fixation and X-gal staining.

Fixation

Whole mounts and cryostat sections were fixed according to published protocols (Liu et al., 1988, and Glaser et al., 1986) specifically designed for X-gal staining of D. melanogaster tissues.

Animals were immersed in freshly prepared 1.0% glutaraldehyde in PBS during dissection and were left in fixative at room temperature for an additional 15 min before a rinse in PBS (modified from Glaser et al., 1986). We found it necessary to dissect the animals thoroughly for adequate fixative penetration. Tissues were fixed and rinsed directly on the Silgard plates.

After air-drying, cryostat sections were fixed in freshly prepared 0.5% glutaraldehyde in PBS for 10 min at room temperature and were rinsed in PBS three times, for 10 min each time (after Liu et al., 1988).

Staining procedure

Whole mounts and cryostat sections were stained with X-gal (protocol after Liu et al., 1988) in a staining buffer; duplicate sets were processed at pH 7.2 - inappropriate for the optimal activity of E. coli [Beta]-galactosidase - and at pH 6.0 - appropriate for the endogenous [Beta]-gal-1 enzyme (Fuerst et al., 1987). Any visible staining of the endogenous [Beta]-galactosidase activity at pH 7.2 can be directly applied as expected background activity in X-gal staining assays designed to detect lacZ fusion gene activity.

In developing our methods, it became apparent that the X-gal should only be dissolved in N,N-dimethyl formamide immediately before staining. Once dissolved in the solvent, the X-gal begins to deteriorate (even when stored at -20 [degrees] C) and staining intensity is reduced. If the age of the X-gal/N,N-dimethyl formamide solution is not constant between different samples, any quantitative measurement (albeit rough) of enzyme activity is compromised.

For comparison of enzyme activity in cryostat sections at the two pH levels, serial sections were collected alternately between two slides. One slide was then stained at pH 7.2 and the other at pH 6.0. This allowed precise comparisons between regions.

After fixation, cryostat sections were encircled with a hydrophobic barrier (Pap Pen, Kiota International Industries), immediately covered with a droplet of freshly prepared X-gal staining solution, and incubated in humidified covered trays for 16 h at 37 [degrees] C. The X-gal solution was removed, and the sections were rinsed three times, 10 min each rinse, in PBS and mounted in Gel-Mount (Biomedia Corp.). Slides were stored in the dark at 4 [degrees] C to preserve staining intensity (Glaser et al., 1986).

After being stained and photographed, sections of the pupal stages were restained in hematoxylin/eosin. These additional stains were helpful in identifying various developing organs during metamorphosis. To prepare the slides for restaining, coverslips were removed by soaking the slides in PBS for up to 4 days at 4 [degrees] C. After a 2-min rinse in water, sections were stained in Harris' hematoxylin (Luna, 1968) for 2 min, rinsed in tap water for 2 min, and rinsed through a graded series of ethanol (70%-90%). Sections were stained in eosin (a saturated solution of eosin in 95% ethanol) for 5 min. Dehydration was continued through 100% ethanol; sections were then cleared in toluene and mounted in Histoclad (Becton, Dickinson and Co.).

Animals dissected and fixed for whole-mount staining were given a final 15-min rinse in PBS and a brief rinse in staining buffer. Dissections were left pinned out on the Silgard plates and were immersed in X-gal staining solution, covered with a petri lid, placed in humidified covered trays, and incubated for 16 h at 37 [degrees] C.

After staining, dissections were rinsed in PBS three times, 15 min each rinse. Organs of interest were removed and photographed as whole mounts. In the dissections stored in PBS in the dark at 4 [degrees] C for a few days, the staining intensity remained the same, but afterwards some stain began to diffuse from intensely stained regions into adjacent nonstained tissues. For this reason, dissections were viewed within 2 days of staining. For long-term storage, whole mounts were stored in 100% glycerol at 4 [degrees] C (Glaser et al., 1986).

Photomicrography

Sections and whole-mount preparations were examined using bright-field microscopy and photographed on a Leitz Dialux 20 microscope fitted with a Wild Photoautomat MPS 55 loaded with Kodak or Fuji 200 ASA color print film.

Transmission electron microscopy

To determine whether microorganisms were indeed eliminated from the gut in axenically reared animals, we first placed intestinal smears on nutrient and EMB agar media (Difco Laboratories). Microorganisms did not grow on these culture plates. We then viewed sections of the larval midintestine that exhibited [Beta]-galactosidase activity under the transmission electron microscope. This ultra-structural examination afforded an additional control: commensals present in the lumen and specialized to that environment might not grow on culture plates, but commensals and mycetocytes, insect cells that incorporate microorganisms (Chapman, 1969), can be distinguished at the ultrastructural level (Smith, 1968).

Stained segments of the larval midintestine were excised and stored temporarily in PBS with Ca[Cl.sub.2] at 4 [degrees] C in plankton-net vials. Tissues were osmicated in a solution of 2% osmium tetroxide in PBS on ice for 1 h, rinsed in distilled water twice, and left in 70% ethanol overnight. Tissues were given two 7-min rinses, in 70% ethanol on ice; rinsed in 95% ethanol, 100% ethanol and propylene oxide three times, 7 min each rinse, at room temperature; and placed in a 1:1 solution of propylene oxide and Epon-Araldite epoxy resin in open vials for overnight infiltration.

Tissues were transferred to a drop of fresh resin and placed in an evacuated cool oven twice, for 20 min each time. Specimens were embedded in fresh resin and cured for 24 h at 60 [degrees] C. Blocks were sectioned on a Sorvall Porter-Blum MT2-B ultramicrotome using glass knives broken on an LKB KnifeMaker. Thick sections cut in the transverse plane through regions of midintestine that stained with X-gal retained a light blue stain on the lumenal surface. To enhance contrast, thick sections were stained with toluidine blue and mounted in Permount (Fisher Scientific).

After screening the thick sections, we cut gold-to-silver ultrathin sections from the blocks. Sections were placed on grids, stained with a saturated solution of uranyl acetate in 100% ethanol for 20-30 min, rinsed with distilled water, stained in a 2% solution of lead citrate for 4 min, and rinsed in distilled water. After the grids were air-dried, sections were viewed on a Phillips CM 10 transmission electron microscope.

Results

Axenic cultures

Axenic conditions did not alter development in D. melanogaster except to slow the timing of larval development. This effect has been noted previously in axenic or mono-axenic cultures (Ashburner and Thompson, 1978). Compared to nonaxenic cultures, it took approximately 24 h longer for axenic cultures to reach the wandering third-instar stage. In all other respects, the axenic cultures appeared healthy and normal; the sterile conditions did not visibly affect the size or appearance of the larval, prepupal, pupal, or adult stages.

Transmission electron microscopy

Transverse sections through regions of the late third-instar larval midintestine were viewed by transmission electron microscopy. These sections of midintestine, including nonstained sections and those exhibiting intense X-gal staining associated with the lumenal surface, did not contain bacteria. The sections were inspected at several magnifications (from 6000 to 15,000X), and there was no evidence of bacterial flora residing in the lumen or housed intracellularly in mycetocytes [ILLUSTRATION FOR FIGURE 1 OMITTED!.

Staining of endogenous [Beta]-galactosidase activity at pH 7.2

Late third-instar larval stages. In the late third-instar larva, X-gal staining of [Beta]-galactosidase activity was found in the anterior and posterior spiracles, larval midintestine, lymph glands, cellular epidermis and in imaginal discs of the eye and antenna [ILLUSTRATION FOR FIGURE 2 OMITTED!. The only difference between the staining patterns of the active-roaming and the later inactive third-instar larvae was in staining intensity.

In active-roaming late third-instar larvae, intense staining was detected in the region of the anterior spiracles, specifically localized to the atria and region of the primordial dorsal prothoracic (or humoral) discs and spiracular glands. Two regions of intense staining were clearly demarcated in the midintestine: one just posterior to the proventriculus and the other just anterior to the junction between the midintestine and hindintestine. Cryostat-frozen and thick-plastic transverse sections through these midintestinal regions showed that the staining was associated with the lumenal surface of the intestinal epithelium. Light staining was found in the larval lymph glands [ILLUSTRATION FOR FIGURE 3 OMITTED] and eye and antenna imaginal discs of some active larvae [ILLUSTRATION FOR FIGURE 4 OMITTED!. There was light staining of the inner cellular epidermis [ILLUSTRATION FOR FIGURE 5 OMITTED].

In inactive late third-instar larvae, which are entering the quiescent stage just preceding pupariation, staining of the eye and antenna imaginal discs [ILLUSTRATION FOR FIGURE 5A OMITTED], lymph glands [ILLUSTRATION FOR FIGURE 5B AND C OMITTED], and cellular epidermis [ILLUSTRATION FOR FIGURE 5D AND E OMITTED! was noticeably more intense than in active third-instar larvae.

Prepupal stages (P1-P4). In white prepupae (P1), whole mounts showed a further increase in the staining intensity of the inner cellular epidermis, eye and antenna discs and lymph glands. Staining in the midintestine persisted. Staining in the now-everted anterior and posterior spiracles, however, was lighter than in late third-instar larvae. Cryostat sections showed new areas of staining: loose, punctate clusters of stained cells distributed throughout the body but not directly associated with any organ, and a light, punctate staining in the anterior organs. As in late third-instar larvae, the wing and leg discs did not stain, nor did the brain, ventral ganglion, or fat body.

In prepupal stage P2, both whole mounts and sectioned material showed that the loose clusters of punctate staining increased in number; staining was also found in the developing eyes and antennae and was now found in the everted legs and wings as well [ILLUSTRATION FOR FIGURE 6A AND B OMITTED!. Clusters of stain appeared in cavities within the evaginated wing rudiments and were found internally at the proximal end of developing legs. The anterior region housing the imaginal discs had areas of foamy-appearing tissue, rich in droplets, probably of lipid, surrounded by diffuse stain. The lymph glands [ILLUSTRATION FOR FIGURE 6C OMITTED] continued to stain intensely, and the midintestine, which at this stage is in the process of being remodeled, now stained more darkly. The brain and ventral ganglion were beginning to change shape from the larval form but did not stain, nor did the fat body.

In prepupal stage P3, the staining patterns evident in P2 persisted and intensified. The punctate clusters of stain became more abundant and were distributed throughout the body. At this stage the abdominal pericardial cells began to show dark staining [ILLUSTRATION FOR FIGURE 6D OMITTED!. The developing brain and ventral ganglion did not stain.

In the prepupal stage P4, the amount of foamy-appearing tissue with diffuse stain increased, and stain was now found throughout the developing intestinal system. The fat body showed light staining. Again, there was no staining in the brain or ventral ganglion.

Pupal stages (PS-P15). During metamorphosis, the delicate nature of the remodeling tissue disallowed whole-mount dissections, but X-gal staining of cryostat serial sections revealed dramatic changes in staining intensity and distribution [ILLUSTRATION FOR FIGURE 6 AND 7 OMITTED].

After head eversion, P5, and in stage P6, there was an increase in dark, punctate staining which co-localized with the fat-body tissue of the head capsule, thorax, and abdomen. By stage P6, dark staining was associated with the first pupal spiracles and trachea. The staining associated with foamy-appearing tissues was no longer found in the head region but persisted in the developing legs and wings. The relatively compact tissues of the developing digestive system showed little staining. The brain and ventral ganglion did not stain.

The highest intensity of staining was apparent in stages P7 and P8 [ILLUSTRATION FOR FIGURE 6E OMITTED]. Dark, punctate staining was distributed throughout the fat body, and a concentrated "ball" of stain appeared at the tip of the abdomen. As with the earlier stages, there was little or no staining of compact tissues. The staining associated with the developing appendages, pupal spiracle, and trachea persisted.

In stage P9, the stain in the thoracic and abdominal fat body decreased and remained at low levels through P14. At stage P10 and in subsequent pupal stages, no stain was found in the developing appendages.

Through stages P9-P14, intense stain was associated with the anterior spiracle region, along the line of the first dorsal anastomosis and into the dorsal tracheal trunks [ILLUSTRATION FOR FIGURE 6F OMITTED] (anatomy after Whitten, 1980). In pupal stages P12-P15, staining at the spiracle was not confined to the tracheal tissues but also appeared as clusters of stain in the tissue surrounding the trachea [ILLUSTRATION FOR FIGURE 6G OMITTED!. These clusters of stain appeared to be the developing thoracic nephrocytes, which stain in the adult. Punctate staining appeared between the muscle masses forming in the dorsal thorax in the region where air sacs are developing. A patch of dark stain also consistently appeared in stages P12-P15 at the dorsal tip of the abdomen in a region where the meconium is found. The meconium, which contains the remains of the larval intestinal epithelium, gastric caecae (Bodenstein, 1965), and other by-products of metamorphosis such as excess hormones (Bownes, 1990), is excreted after eclosion.

Adults. In dissected adult flies, intense X-gal staining was found in the thoracic nephrocytes [ILLUSTRATION FOR FIGURE 8A OMITTED], abdominal pericardial cells [ILLUSTRATION FOR FIGURE 8B OMITTED], and two clearly demarcated regions of the ventriculus [ILLUSTRATION FOR FIGURE 8C OMITTED!. Staining was also consistently found in the reproductive systems of male and female flies aged 1 to 2 days, 3 to 5 days, and 7 to 10 days. There was staining in the ejaculatory pump of 1- to 2-day-old male flies [ILLUSTRATION FOR FIGURE 8D OMITTED!, which intensified in the older 3- to 5- and 7- to 10-day-olds [ILLUSTRATION FOR FIGURE 8C AND 9 OMITTED]. In the female, small clusters of stain were found in the calyx and pedicel of the ovary [ILLUSTRATION FOR FIGURE 8E AND 9 OMITTED], and there was dark staining of the uterus [ILLUSTRATION FOR FIGURE 8F AND 9 OMITTED]. In newly eclosed flies, punctate staining was found associated with the fat body and there was diffuse light staining of the rectal papillae. In the reproductive systems of immature females, staining was found only in the uterus.

Staining of endogenous [Beta]-galactosidase activity at pH 6.0

Third instar larval and prepupal stages (Table I). In the third-instar larval and prepupal stages, the staining patterns at pH 6.0 differed from those at pH 7.2 primarily in the intensity of staining. Areas that stained at pH 7.2 (cellular epidermis, eye-antenna disc, everting legs and wings, lymph glands, midintestine, larval spiracles, internal punctate staining regions, and pericardial cells) stained more intensely at pH 6.0. The pericardial cells stained at an earlier stage at pH 6.0 than at 7.2. Some structures that stained at pH 6.0 did not stain at pH 7.2: the garland (wreath) cells, which stained darkly, and the proventriculus, ventral ganglion, and testes - all of which stained lightly.

At pH 6.0, as at pH 7.2, overall staining levels increased as the animals approached pupation.

Pupal stages (Table II). In the pupa, as in the prepupal stages, the primary difference in staining patterns between specimens stained at the two pH levels was in intensity of staining. Areas that stained at pH 7.2 (fat body, developing legs and wings, first pupal spiracle, tracheae, punctate staining between dorsal thoracic muscles, and developing digestive system) stained more intensely at pH 6.0. Staining persisted longer in developing legs and wings at pH 6.0 than at pH 7.2. Some structures in cryostat sections stained at pH 6.0 but not at pH 7.2: brain and ventral ganglion and antennae.

As at pH 7.2, the intensity of staining increased with increased age between stages P4 and P8. The intensity appeared to peak at P7-P9.

Adult stages (Table III). In the adult, as in the other stages, a major difference between staining patterns at pH 7.2 and 6.0 was in intensity, with areas that stained at pH 7.2 staining more intensely at pH 6.0 (ventriculus, fat body, calyx and pedicel, and sperm pump). Some regions of staining did not change in intensity (thoracic nephrocytes and pericardial cells). Some structures that stained at pH 6.0 did not stain at pH 7.2: testes, antennae and brain and, in the newly enclosed flies, the thoracic ganglion and halteres. The staining in the brain was pronounced [TABULAR DATA FOR TABLE I OMITTED! in newly enclosed flies, including areas of the cellular cortex, cell body layer of the lamina, and the first and second optic chiasmata [ILLUSTRATION FOR FIGURE 10 OMITTED]. In adults aged 1-2 days and older, staining persisted only in the cell-body layer of the lamina. No differences in these staining patterns were noted between males and females.

Discussion

We have characterized tissue-specific activity of endogenous [Beta]-galactosidase in wild-type Drosophila melanogaster by staining with the chromogen X-gal at pH 7.2 and 6.0. Our findings show that at both pH levels, [Beta]-galactosidase activity is found in a variety of tissues in the larval, pupal, and adult stages and that striking changes in tissue-specific activity of the enzyme occur at the larval-pupal transition and through metamorphosis. The staining patterns shown at pH 7.2 are particularly germane for studies in which X-gal is used to assay lacZ fusion gene or enhancer trap activity in D. melanogaster. At the staining pH typically recommended for these studies (pH 7.2; Ashburner, 1989), animals will not only exhibit endogenous enzyme activity in certain tissues throughout developmental stages, but will also exhibit sites of activity that change within a single stage.

In our study, we discovered certain tissue-specific activities that have not previously been reported. At staining pH levels of 7.2 and 6.0, sites of activity were the lymph glands, cellular epidermis, and eye-antenna imaginal discs of the larva and the thoracic nephrocytes and fat body of the adult. At a staining pH of 6.0, additional sites were the larval garland (wreath) cells, proventriculus, ventral ganglion and gonads; the pupal brain, ventral ganglion, and antennae; and the adult brain and antennae, with the halteres and thoracic ganglion staining in the newly enclosed fly. These results, coupled with the fact that staining at pH 6.0 in general was more intense than at pH 7.2, show that staining patterns are pH-dependent.

A previous report of endogenous [Beta]-galactosidase activity in the larval midintestine, adult ventriculus, pericardial cells, and ejaculatory pump (Glaser et al., 1986) was based on cultures that were not grown axenically. Because our axenic cultures showed the same activity, we conclude that these regions of [Beta]-galactosidase activity are not due to the presence of commensal flora in the fly. In limited studies of nonaxenic cultures, we did, however, find instances of variable X-gal staining that we did not find in the axenic cultures; for example, in the adult (3-day and older) appendages and rectum (unpubl. data). This suggests that bacterial contamination can contribute to the X-gal staining reaction, and that it may be wise to grow flies axenically to eliminate this variable.

Our results should not preclude the use of transformed flies in a lacZ/X-gal assay, but the pH of the staining buffer should be carefully monitored and the flies should be staged precisely. If staging errors are made - for example, if an active third-instar larva is used as a control animal but a slightly older inactive transformed larva is assayed for lacZ fusion-gene activity - the pronounced endogenous activity in the eye and antenna imaginal discs, lymph glands, and cellular epidermis could be construed as ectopic expression of the lacZ fusion-gene product. In some reports from studies on D. melanogaster using lacZ fusion gene transformants, X-gal staining in the eye and antenna discs of third-instar larvae was attributed to the fusion-gene product, with no mention of any similar staining patterns in the controls (e.g., Glaser et al., 1986; Brand and Perrimon, 1993). In light of our evidence for the need to control for age-dependent expression within the same life stage, such results should be reexamined with attention to the staging of the larvae. In addition, the finding that, during metamorphosis, the endogenous enzyme activity is expressed at high levels in many tissues - such as [TABULAR DATA FOR TABLE II OMITTED] the developing legs, wings, alimentary tract, and fat body - even at a staining pH (7.2) that is meant to exclude endogenous activity, use of an anti-E. coli [Beta]-galactosidase antibody, a different reporter gene (such as luciferase), or a different molecular assay would be appropriate when investigating gene activity in these tissues.

The presence of [Beta]-galactosidase activity in several organ systems and throughout metamorphosis suggests that this enzyme might be involved in several vital physiological processes in D. melanogaster. The increases in the distribution and level of enzyme activity before and during metamorphosis are particularly interesting because they coincide with known peaks of ecdysteroid titer (Bainbridge and Bownes, 1988; Pak and Gilbert, 1987; and Handler, 1982). The appearance of endogenous [Beta]-galactosidase activity in the ventral ganglion, eye and antenna imaginal discs, lymph glands, and cellular epidermis in late third-instar larvae just prior to pupariation coincides with a dramatic increase in ecdysteroid titer 10 h before puparium formation (Bainbridge and Bownes, 1988). [TABULAR DATA FOR TABLE III OMITTED! Additionally, the increase in [Beta]-galactosidase activity in pupal stages P5-P9, peaking at stages P7 and P8, coincides with a high concentration of ecdysteroids that peaks at P7 (Bainbridge and Bownes, 1988). The possibility of a relationship between ecdysterone and [Beta]-galactosidase expression is further supported by the observation that ecdysterone induces the expression of [Beta]-galactosidase in D. melanogaster cell cultures (Best-Belpomme et al., 1978).

The physiological role of [Beta]-galactosidase in D. melanogaster has not been characterized. If the [Beta]-galactosidase in D. melanogaster is involved in activities akin to those of a mammalian lysosomal form (for example, see Wallenfels and Weil, 1972; Belfiore, 1980), the induction and expression of the enzyme during metamorphosis is not surprising, because enzymatic degradation of macromolecules is widespread during this period. During metamorphosis, larval cells undergo histolysis and are attacked by phagocytic cells. These phagocytic cells degrade the debris from the larval cells, and material is recycled for use by the developing imaginal tissues (Chapman, 1969). Many of the areas that we found to have [Beta]-galactosidase activity are associated with phagocytic activity: the lymph glands release large numbers of phagocytic cells into the haemocoel through the prepupal stage (Rizki, 1978a); in the eye and antenna discs, localized programmed cell death, correlated with high concentrations of lysosomes (Murphy, 1974), occurs during morphogenesis (Nothiger, 1972; Poodry, 1980; Kankel et al., 1980); the protein globules of the fat body are thought to be cytolysosomal or autophagic bodies (Rizki, 1978b) that are discharged from the cells into the intercellular space during pupation (Yagi, 1962); the garland cells, thoracic nephrocytes, and pericardial cells, considered stationary macrophages, engulf microbes and remove toxins from the haemolymph - a role analogous to that of the vertebrate reticuloendothelial system (Wigglesworth, 1972). To determine whether the [Beta]-galactosidase activity that we see is found in lysosomal compartments, we plan to examine the sub-cellular localization of the enzyme in the lymph glands, thoracic nephrocytes, and pericardial cells.

The physiological significance of invertebrate [Beta]-galactosidases is unexplored. In light of our findings, D. melanogaster appears to be an appropriate organism for such an investigation.

Acknowledgments

We thank Drs. John Ringo, Harold Dowse, Irv Kornfield, and Alex Parker for their critique of this work. The work was supported in part by grants to H. Dowse from the American Heart Association and the National Institutes of Health (grant GEN RO1 NS26412) and by a Faculty Research Fund Award to M. Tyler. We are grateful for this support.

Literature Cited

Ashburner, M. 1989. Drosophila: A Laboratory Handbook. Cold Spring Harbor Press, Cold Spring Harbor, NY.

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Author:Schnetzer, Jamie Watler; Tyler, Mary S.
Publication:The Biological Bulletin
Date:Apr 1, 1996
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